Interactions between biosynthesis, compartmentation and transport

Transcription

Journal of Experimental Botany, Vol. 53, No. 372,
Antioxidants and Reactive Oxygen Species in Plants Special Issue,
pp. 1283–1304, May 2002
Interactions between biosynthesis, compartmentation
and transport in the control of glutathione homeostasis
and signalling
Graham Noctor1, Leonardo Gomez2, Hélène Vanacker3 and Christine H. Foyer 3,4
1
Institut de Biotechnologie des Plantes, Bât 630 Université Paris VII, 91405 Orsay Cedex, France
Instituto de Fitopatologia y Fisiologia Vegetal, CNIA-INTA, Cno. 60 Cuadras Km 51u2, X5020ICA
Cordova, Argentina
3
Crop Performance and Improvement, IACR-Rothamsted, Harpenden, Herts AL5 2JQ, UK
2
Received 10 July 2001; Accepted 10 December 2001
Abstract
Glutathione has numerous roles in cellular defence
and in sulphur metabolism. These functions depend
or impact on the concentration anduor redox state
of leaf glutathione pools. Effective function requires
homeostatic control of concentration and redox state,
with departures from homeostasis acting as signals
that trigger adaptive responses. Intercellular and
intracellular glutathione pools are linked by transport
across membranes. It is shown that glutathione can
cross the chloroplast envelope at rates similar to the
speed of biosynthesis. Control of glutathione concentration and redox state is therefore due to a complex
interplay between biosynthesis, utilization, degradation, oxidationureduction, and transport. All these
factors must be considered in order to evaluate the
significance of glutathione as a signalling component during developement, abiotic stress, or pathogen
attack.
Key words: Chloroplasts, c-glutamylcysteine synthetase,
glutathione, maize, plant–pathogen interaction, transporter,
Triticum aestivum, wheat, Zea mays.
Introduction
Glutathione (c-Glu-Cys-Gly) is a multifunctional metabolite in plants (Fig. 1). It is a major reservoir of
4
non-protein reduced sulphur, and has crucial functions in
cellular defence and protection. Glutathione reacts chemically with a range of active oxygen species (AOS), while
enzyme-catalysed reactions link GSH to the detoxification of H2O2 in the ascorbate–glutathione cycle. Importantly, GSH protects proteins against the denaturation
that is caused by oxidation of protein thiol groups during
stress. All these functions involve the oxidation of the
thiol group, principally to form glutathione disulphide
(GSSG). Cellular GSH : GSSG ratios are maintained by
glutathione reductase (GR), a homodimeric flavoprotein
that uses NADPH to reduce GSSG to two GSH. Like all
other aerobic organisms, plants maintain cytoplasmic
thiols in the reduced (–SH) state because of the low thioldisulphide redox potential imposed by millimolar amounts
of glutathione, which acts, therefore, as a thiol buffer.
Although transient disulphide bonds do occur during the
catalytic cycle of some enzymes, stable protein disulphide
bonds are relatively rare except in quiescent tissues such
as seeds. The multiple roles of GSH within the cell,
together with the stability of GSSG, may make this redox
couple ideally suited to information transduction. The
GSHuGSSG ratio is likely to be far more influential in
the control of gene expression and protein function than
the absolute size of the glutathione pool. This review
specifically addresses the key factors involved in glutathione homeostasis (synthesis, sinks, partitioning, and
transport) that are central to the putative roles of this
compound in signalling.
To whom correspondence should be addressed. Fax: [ 44 (0)1582 763010. E-mail: [email protected]
Abbreviations: AOS, active oxygen species; CAT, catalase; chl chlorophyll; CHS, chalcone synthase; DHA, dehydroascorbate; DHAR,
dehydroascorbate reductase; c-EC, c-glutamylcysteine; c-ECS, c-glutamylcysteine synthetase; FW, fresh weight; GPX, glutathione peroxidase;
GR, glutathione reductase; GSH, reduced glutathione; GSH-S, glutathione synthetase; GSSG, glutathione disulphide; GST, glutathione S-transferase;
JA, jasmonic acid; PGA, 3-phosphoglyceric acid; SA, salicylic acid; SAR, systemic acquired resistance.
ß Society for Experimental Biology 2002
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Noctor et al.
Fig. 1. Glutathione biosynthesis and interacting processes in plant cells. c-EC, c-glutamylcysteine; c-ECS, c-glutamylcysteine synthetase; GR,
glutathione reductase; GSH, reduced glutathione; GSSG, glutathione disulphide; GSH-S, glutathione synthetase; GST, glutathione S-transferase.
In addition to its antioxidant functions, glutathione
is a precursor of phytochelatins and a substrate for
glutathione-S-transferase (GST: Fig. 1). Over and above
its function in the ascorbate–glutathione cycle, glutathione acts as a direct electron donor to peroxides in
reactions catalysed by glutathione peroxidase (GPX). The
animal GPX is a selenoprotein that detoxifies H2O2 at
high rates. By contrast, plant GPXs are not constitutive
but are induced in response to stress. They do not contain
selenium and only catalyse GSH-dependent reduction of
H2O2 at rates which are very low compared with the high
rates of H2O2 generation in plants (Foyer and Noctor,
2000). In plants, ascorbate peroxidase (APX) and catalase
(CAT) are predominant in the detoxification of H2O2
while GPXs are more important in other areas of oxidant
metabolism, including the removal of lipid and alkyl
peroxides (Eshdat et al., 1997). The only clear demonstration of GPX targeting thus far has shown direction to
the chloroplast (Mullineaux et al., 1998). In addition to
their role in conjugation, GSTs can use GSH to reduce
peroxides (Cummins et al., 1999). Transgenic tobacco
lines overexpressing plant GSTuGPX were reported to
show enhanced antioxidant capacity and substantial
improvement in seed germination and seedling growth
under stress (Roxas et al., 1997). GSTs form a large,
heterogeneous family of proteins that share the defining
characteristic of catalysing the nucleophilic attack of the
sulphur atom of GSH (or homologue) on the electrophilic
centre of their substrates. They are therefore responsible
for the removal of compounds that are potentially genotoxic or cytotoxic by virtue of their reaction with
electrophilic sites in DNA, RNA and proteins. It has
become clear, however, that the function of GSTs is
not limited to these reactions: GSTs also seem to be
involved in a ‘ligandin’ function important, for example,
in the anthocyanin synthesis pathway (Marrs, 1996). It
may be that certain GSTs function as flavonoid-binding
proteins as suggested recently for AN9, a GST required
for efficient anthocyanin export from the cytosol in
petunia (Mueller et al. 2000). Consistent with this is the
Glutathione homeostasis and signalling
observation that the anthocyanin content of Arabidopsis
leaves correlated with GSH content in plants with
modified capacity for GSH biosynthesis ( Xiang et al.,
2001).
While it has long been accepted that glutathione is
essential for vigour, it has only recently been recognized
that this tripeptide cannot be functionally replaced,
except perhaps by one of its homologues. The rml1
mutant of Arabidopsis, which is deficient in c-ECS and
contains no detectable glutathione, has a marked phenotype with an absence of root development and a small
shoot system, and can survive only in tissue culture supplied with GSH (May et al., 1998a). Similarly, transgenic
Arabidopsis with less than 5% wild-type leaf glutathione
contents were shown to be significantly decreased in size
and biomass, and were more sensitive to environmental
stress (Xiang et al., 2001). A strong correlation has also
been demonstrated beween root GSH content and the
capacity of the cells in the root apical meristem to
proliferate (May et al., 1998a). However, although a
96% reduction of shoot glutathione contents was associated with shorter roots in transformed Arabidopsis, the
decrease in root length was only of the order of 40%
(Xiang et al., 2001).
All these recent developments underline the importance of the control of glutathione concentration and
redox state in plant cells. This review discusses the many
advances made over the last decade in the understanding
of glutathione biosynthesis, and also begins to address
the key issue of compartmentation and related transport
processes. These processes, complex and poorly defined,
are discussed here in the light of new data obtained in the
authors’ laboratory.
Biosynthesis of glutathione
Enzymes and genes
The pathway of glutathione biosynthesis is well established and is similar in plants, animals and microorganisms. In two ATP-dependent steps, catalysed by
c-glutamylcysteine synthetase (c-ECS) and glutathione
synthetase (GSH-S), the constituent amino acids are
linked to form the complete tripeptide (Fig. 1). The
N-terminal peptide bond linking glutamic acid to
cysteine in GSH is unusual in that glutamic acid is
linked via the c- rather than the a-carboxyl group. The
low activities of c-ECS and GSH-S in plants, and the
complexities of the procedures for enzyme extraction and
assay, have precluded extensive purification and kinetic
characterization. Consequently, much of the current
knowledge of their structure, regulation and function has
been gleaned from molecular techniques and plant transformation. It is clear that the two-step reaction sequence
occurs in both chloroplastic and non-chloroplastic
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compartments and is found in photosynthetic and
non-photosynthetic tissues (Foyer and Noctor, 2001).
A gene encoding c-ECS, here denoted as gsh1, was
originally cloned from Arabidopsis thaliana by complementation of an E. coli mutant deficient in this
enzyme (May and Leaver, 1994). Heterologous expression of the Arabidopsis c-ECS in a yeast mutant recovered
only 10% of the GSH measured in the wild-type yeast
(May and Leaver, 1994). This discrepancy provoked
much speculation concerning the identity of the cloned
gene, but further complementation studies have now
confirmed that this gene does indeed encode a protein
with true c-ECS activity (May et al., 1998a).
Functional complementation of an E. coli mutant
deficient in GSH-S activity was also used to clone the
Arabidopsis thaliana gene for this enzyme, which is
denoted here as gsh2 (Rawlins et al., 1995). The ability
of several plant species to make homologues of
glutathione depends on the specificity of the synthetases
involved. Specific legume GSH-Ss use either glycine to
form GSH or b-alanine to form homoglutathione. Recent
work in Medicago truncatula suggests that separate
genes encode GSH-S and homoglutathione synthetase
(hGSH-S) and that the divergence in specificity has arisen
by gene duplication after the evolutionary divergence
of the Leguminosae (Frendo et al., 1999). The two genes
are very homologous and are found on the same fragment
of genomic DNA. In a consideration of the distribution
of the biosynthetic enzymes in legume nodules, Becana
et al. have suggested that c-ECS is plastidic, hGSH-S is
cytosolic and GSH-S isoforms exist in both the cytosol
and mitochondria in several legume species (Becana et al.,
2000).
Regulation of biosynthesis
Glutathione synthesis is controlled primarily by c-ECS
activity and cysteine availability: As in animals, the
activity of c-ECS limits the rate of glutathione synthesis in
plants under most conditions. Consistent with this notion
is the observation that the cad-2 Arabidopsis mutant,
which has a mutation in the gsh1 gene, has only one-third
of the tissue glutathione contents of the wild-type
(Cobbett et al., 1998). Antisense suppression of c-ECS
in Arabidopsis also causes substantial decreases in leaf
glutathione ( Xiang et al., 2001). A key role for c-ECS
in controlling the rate of glutathione synthesis is supported by the increases in extractable enzyme activity in
tissues treated with cadmium (Rüegsegger and Brunold,
1992). Most tellingly, overexpression of an E. coli c-ECS
but not GSH-S, in poplar or tobacco substantially
increases leaf glutathione contents (Strohm et al., 1995;
Noctor et al., 1996, 1998; Creissen et al., 1999), as does
homologous overexpression of the Arabidopsis c-ECS
(Xiang et al., 2001).
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Noctor et al.
Bacterial genes encoding c-ECS and GSH-S have been
introduced into poplar, mustard and tobacco (Strohm
et al., 1995; Foyer et al., 1995; Noctor et al., 1996, 1998;
Zhu et al., 1999; Pilon-Smits et al., 1999; Creissen et al.,
1999). Over-expression, with targeting of the bacterial
enzyme protein to either the chloroplast or cytosol, led
to marked increases in enzyme activity. Increases in
c-ECS, but not GSH-S, not only led to constitutive
increases in leaf glutathione (up to 400%: Noctor et al.,
1996, 1998; Creissen et al., 1999) but glutathione was
also increased in xylem sap, phloem exudates and roots
(Herschbach et al., 2000). Of particular note is the
observation that the cysteine pool was not depleted by
the increased demand for thiols, but was even slightly
enhanced in response to increased c-ECS activities,
pointing to co-ordinate regulation of cysteine synthesis
and glutathione synthesis. This observation is supported
by leaf thiol contents in plants homologously overexpressing the plant c-ECS (Xiang et al., 2001). Despite
enhanced GSH contents in the phloem of poplar, sulphur
uptake by the roots was markedly enhanced to meet
the requirements of increased demand for sulphur
(Herschbach et al., 2000). Nevertheless, incubation of
leaf discs with cysteine increased glutathione contents
substantially in untransformed and transformed poplars,
particularly in the light, suggesting that cysteine supply
remains a key limitation (Strohm et al., 1995; Noctor
et al., 1996, 1997).
The dipeptide produced by the c-ECS reaction is present at very low levels in most untransformed plants. In
the poplars overexpressing c-ECS, however, c-EC was
greatly increased. In some conditions, this was attributable to insufficient availability of Gly (Noctor et al.,
1997). Even when Gly was abundant, however, c-EC was
still much higher than in untransformed plants, reflecting
a shift in control from c-ECS to GSH-S, whether the
bacterial c-ECS was present in the cytosol or chloroplast
(Noctor et al., 1998). This suggested that overexpression
of both enzymes together would increase the potential for
constitutive enhancement of tissue glutathione contents
even further than that achieved by c-ECS overexpression
alone. This effect was observed when tobacco lines
overexpressing each of the biosynthetic enzymes from
E. coli were crossed to produce hybrids over-producing
both enzymes although, suprisingly, c-EC contents were
found to be higher in the hybrid lines than in those lines
overexpressing c-ECS alone (Creissen et al., 1999). The
marked phenotype produced by chloroplastic c-ECS
overexpression in tobacco complicates the interpretation
of these results. By contrast, a phenotype linked to
chloroplastic c-ECS overexpression was not observed in
poplar ( Noctor et al., 1998) or Brassica juncea (Zhu et al.,
1999; Pilon-Smits et al., 1999), except that these transformed plants were more, rather than less, stress tolerant. Similar results have been reported in transformed
Arabidopsis overexpressing the endogenous c-ECS ( Xiang
et al., 2001). The above evidence demonstrates that the
most important factors controlling plant glutathione are
the activity of c-ECS and the availability of cysteine,
and recent work suggests that these two factors may
be co-ordinated (H Rennenberg, personal communication). The in vivo activity of c-ECS is determined by
control at multiple levels, and these are discussed in the
following sections.
Control of transcription and translation of c-ECS and
GSH-S: Studies in animals, particularly on cancer cells
challenged with chemotherapeutic agents, have shown
that transcription of the c-ECS gene is regulated by
protein factors and by conserved antioxidant response
elements upstream of the coding sequence (Foyer and
Noctor, 2001). Relatively little is known about the
co-ordinate regulation of expression of gsh1 and gsh2 in
plants, but it is clear that GSH and GSSG per se exercise
little or no control over transcription ( Xiang and Oliver,
1998). Similarly, H2O2 did not affect transcript abundance. The abundance of gsh1 and gsh2 transcripts was
increased by cadmium in Brassica juncea (Schäfer et al.,
1998) and by both cadmium and copper in Arabidopsis
( Xiang and Oliver, 1998). Jasmonic acid (JA) also
increased gsh1 and gsh2 transcripts and a common
signal transduction pathway may be involved (Xiang
and Oliver, 1998). Interestingly, JA is involved in the
control of glucosinolate synthesis, activation of which
can represent a significant increase in sulphur demand in
the Brassiceae (Doughty et al., 1995). Although transcript
abundance was increased by heavy metals and JA, oxidative stress was required for the translation of the transcripts, implicating regulation at the post-transcriptional
level and a possible role for factors such as H2O2 or
modified GSHuGSSG ratios in de-repressing translation
of the existing mRNA (Xiang and Oliver, 1998). It is
interesting to note that among the most spectacular
increases in glutathione are those observed when plants
deficient in CAT are placed in conditions favouring
photorespiration (Smith et al., 1984; Willekens et al.,
1997), where the accumulation in total glutathione is
accompanied by a marked decrease in the reduction state
of the pool. A similar response was elicited by exposing
poplar leaves to ozone (Sen Gupta et al., 1991). The
59untranslated region (59UTR) of the gsh1 gene was
found to interact with a repressor-binding protein that
was released upon addition of H2O2 or changes in the
GSHuGSSG ratio (Xiang and Bertrand, 2000). A redoxsensitive 59UTR-binding complex is thus suggested to
control c-ECS mRNA translation in A. thaliana ( Xiang
and Bertrand, 2000).
Post-translational control of c-ECS: Post-translational
regulation of c-ECS through end-product inhibition by
GSH is a crucial factor in controlling GSH concentration in animals and plants. Covalent modification
may also be influential. There is some evidence to suggest
that rat c-ECS is regulated by protein phosphorylation
(Sun et al., 1996) but this has not yet been found in
studies on c-ECS from plants. May et al. concluded
that protein factors are involved in post-translational
control of c-ECS and are required for full activity
(May et al., 1998b). The failure of the plant enzyme to
operate ectopically was explained by the absence of such
endogenous plant factors (May et al., 1998a, b). In the
animal enzyme system, a smaller regulatory subunit acts
to increase the catalytic potential of the larger catalytic
subunit by increasing its Ki value for GSH and decreasing the Km for glutamate, thereby alleviating feedback
control and allowing the enzyme to operate under in vivo
conditions (Huang et al., 1993). It is nevertheless clear
that even in the absence of the smaller subunit, the large
catalytic subunit is capable of effective catalysis, since
overexpression of this polypeptide alone yielded increased
glutathione levels in transfected human cells (Mulcahy
et al., 1995). The highest glutathione contents were, however, obtained by dual overexpression of both subunits
(Mulcahy et al., 1995).
While protein factors have not been identified in plants,
and there is as yet no evidence for control of c-ECS
by phosphorylation, several enzymes in plants are controlled by interactions between phosphorylation status
and factors such as 14-3-3 proteins, regulatory components found in several compartments of the plant cell
(DeLille et al., 2001). This interaction inactivates enzymes
such as nitrate reductase, but also confers stability against
proteolytic attack. It is as yet unclear whether this type of
regulation might be important in glutathione synthesis
in plants, but it is perhaps worth considering a potential
role in the post-translational control of glutathione synthesis. Non-linearity of the specific activity of c-ECS with
protein concentration has been alluded to previously
(Hell and Bergmann, 1990). As part of a study on the
intracellular distribution of GSH metabolism in wheat
(see below), c-ECS activity was measured in unfractionated leaf extracts and in purified chloroplasts. In both
cases, specific activity increased substantially as protein
was increased (Fig. 2). At the highest amounts of protein
in the assay, the activity in wheat leaf extracts was very
similar to that reported previously (approximately
0.5 nmol mg Y 1 protein min Y 1: cf. Fig. 2 and Hell and
Bergmann, 1990). In chloroplast extracts, the activity
showed an almost linear increase with protein and, at
the highest protein concentration, attained values that
were more than 4-fold higher than those found in whole
leaf extracts (Fig. 2). On a chlorophyll basis, maximal
c-ECS activity was approximately twice as high in
chloroplast extracts as in whole leaf extracts. These
effects are unlikely to be due to low-molecular-weight
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effectors since similar results were obtained whether or
not extracts were desalted prior to assay.
c-ECS from whole tobacco and parsley cells has been
shown to be subject to inhibition by GSH, in a manner
that is competitive with respect to glutamate (Hell and
Bergmann, 1990; Schneider and Bergmann, 1995). The
KiGSH was 0.27–0.45 mM at a glutamate concentration of
10–20 mM (Hell and Bergmann, 1990). Figure 3 shows
Fig. 2. The specific activity of c-ECS increases with increasing protein
concentration in the assay. Activity was measured under anaerobic
conditions (adapted from Hell and Bergmann, 1990). Intact wheat
chloroplasts were isolated as described in Table 2, and lysed osmotically
(1 : 5 dilution) into 10 mM HEPES, ( pH 8.0), 5 mM MgCl2, and 1 mM
EDTA. Following vigorous mixing and 20 min incubation on ice,
membranes were removed by centrifugation and 0.23 ml soluble extract
added to 0.26 ml assay buffer in a closed glass HPLC autosampler vial.
Helium was passed though the mix for 10 min to remove oxygen, the
mix was equilibrated at 30 C for 10 min and the reaction was started by
addition of 10 ml de-oxygenated cysteine through the PTFE septum,
using a Hamilton syringe. The final concentration of the assay mix (final
volume 0.5 ml) was 0.1 M HEPES ( pH 8.0), 50 mM MgCl2, 20 mM
glutamate, 5 mM ATP, 5 mM phosphocreatine, 5 U phosphocreatine
kinase, 50 mM glucose, 10 U glucose oxidase, and 100 U catalase.
Samples were withdrawn with a Hamilton syringe after 5, 65 and
125 min, and transferred to an Eppendorf tube containing 0.2 ml
50 mM CHES ( pH 8.5). Monobromobimane (20 ml 30 mM) was added
and the derivatization of thiols stopped after 15 min by addition of 1 ml
10% acetic acid. The mix was centrifuged twice for 30 min at 4 C and
20 000 g. The clear supernatant was filtered and introduced into
autosampler vials. Monobromobimane derivatives of Cys and c-EC
were separated by HPLC and quantified by fluorimetric detection, with
reference to known standard concentrations. The formation of c-EC was
linear between 5 min and 125 min at all protein concentrations. Rates
were obtained by subtracting the amount of c-EC formed after 65 min
and 125 min from that present after 5 min. Loss of Cys not attributable
to c-EC formation was less than 10% after 125 min incubation.
Fig. 3. Inhibition of chloroplastic c-ECS activity by glutathione in the
assay. Methods as in Fig. 2 (glutamate concentration was 20 mM in all
assays). Extracts and inhibitor were co-incubated for 15 min prior to
initiation of the reaction.
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Noctor et al.
Table 1. c-ECS activity extracted from purified chloroplast shows
similar sensitivity to inhibitors to the activity from whole leaves
Activities are given in nmol mg Y 1 protein min Y 1 (% activity in the
absence of inhibitors in brackets). Extracts and inhibitor were
co-incubated for 15 min prior to the initiation of the reaction.
Chloroplasts extracted as in Table 2, c-ECS assayed as in Fig. 2.
The ratio of soluble protein to chlorophyll in the isolated chloroplasts
was approximately half that of whole leaf extracts. BSO, buthionine
sulphoximine.
Inhibitor
No addition
1 mM GSH
10 mM GSH
1 mM BSO
Enzyme activity ( % control)
Leaves
Chloroplasts
0.58
0.54
0.14
0.11
1.07
0.90
0.03
0.14
(100)
(93)
(24)
(18)
(100)
(84)
(3)
(13)
that the enzyme extracted from purified wheat chloroplasts, assayed at a constant protein concentration, was
also sensitive to GSH. c-ECS from wheat chloroplast
and whole leaf material showed a similar sensitivity to
inhibition by GSH and by buthionine sulphoximine
(Table 1). It would appear, therefore, that chloroplastic
and extrachloroplastic isoforms possess similar regulatory
properties, although this notion can only be definitely
confirmed by biochemical studies of the extrachloroplastic enzyme(s). Given the likely difficulties of isolating
cytosolic c-ECS through classical techniques, purification
through heterologous expression of a cloned cDNA offers
the best approach to this question.
Intercellular compartmentation of glutathione synthesis
The implications of heterogeneity of cells with regard to
glutathione metabolism have only recently become the
focus of research effort. Apart from the pioneering
studies of Rennenberg (see below), little information on
this question had appeared in the literature. Given that
the sensitivity of certain cells may be explained by their
lack of adequate glutathione production or recycling, this
question is key.
Compartmentation in C3 leaves: Glutathione is not
produced at equivalent rates by all tissues or, indeed, by
all cells within a tissue. Of particular note is the high
capacity for GSH biosynthesis in some types of trichomes. These specialized unicellular or multicellular
structures on the epidermis can have a protective function
in excreting toxic compounds such as cadmium. The
trichomes found on the stem and leaf surface of various
A. thaliana ecotypes, show much higher expression of
enzymes involved in the synthesis of cysteine and GSH
and have GSH contents 2–3 times higher than the
surrounding basal and epidermal cells (Gutierrez-Alcala
et al., 2000). The evidence for GSH, GSSG and
GS-conjugate transport systems on the plasma membrane
associated with systemic transport has recently been
discussed (Foyer et al., 2001) and hence these systems
will not be elaborated upon here further.
Compartmentation in maize leaves: An extreme example
of the differential intercellular partitioning of glutathione
metabolism is observed in maize. In common with other
plants that show ‘Kranz anatomy’ and C4 photosynthesis,
maize leaves have two photosynthetic cell types whose
functions are very different. The enzymes of the Benson–
Calvin cycle, which are very sensitive to redox regulation,
are localized in bundle sheath chloroplasts. In contrast
to the mesophyll, the bundle sheath cells have very low
amounts of photosystem II, ferredoxin and ferredoxinNADP [ reductase and, therefore, a lower capacity for
the photochemical production of reducing power. H2O2
was found only in the mesophyll compartment in optimal
growth conditions (Doulis et al., 1997) but accumulated
in the bundle sheath cells at low temperatures (Pastori
et al., 2000a). Following exposure to cold stress, oxidative
damage was found almost exclusively in the bundle
sheath (Kingston-Smith and Foyer, 2000).
Recent evidence suggests that the sensitivity of maize
leaves to chilling-induced precocious senescence is related
to the ability to synthesize and regenerate GSH (Kocsy
et al., 2000) and to the absence of antioxidant generation and recycling capacity in the bundle sheath cells
(Kingston-Smith and Foyer, 2000). Like DHAR, GR is
localized only in the leaf mesophyll cells. By contrast,
other antioxidant enzymes are either restricted to the
bundle sheath cells (APX and superoxide dismutase)
or are found to be approximately equally distributed
between the two cell types (CAT and MDHAR; Doulis
et al., 1997; Pastori et al., 2000a). The exclusive localization of GR activity in the mesophyll cells may
be explained by the comparative lack of reductant in
the bundle sheath cells (Doulis et al., 1997). Because of
their low water-splitting capacity, bundle sheath cells may
not generate sufficient NADPH to support the reduction
of GSSG and DHA. GSSG and DHA produced in the
bundle sheath tissues must, therefore, be transported
to the mesophyll tissues to be reduced.
The absence of GR from the maize bundle sheath is due
to post-transcriptional regulation, since GR transcripts
are found in both cell types (Pastori et al., 2000b).
Cysteine is synthesized in the bundle sheath whereas
GSH-S activity is located predominantly in the mesophyll
cells (Burgener et al., 1998). It appears, therefore, that
glutathione is synthesized in the cells where GR is present
and that the bundle sheath relies on the mesophyll for
both the synthesis of glutathione and the reduction
of GSSG. cDNAs corresponding to maize c-ECS and
GSH-S mRNA have recently been isolated. The 1664 bp
nucleotide sequence obtained for c-ECS mRNA
(EMBL Nucleotide Sequence Database Acc. No.
AJ302783) consists of a 38 nt 59 untranslated region
that precedes the first ATG, a 1317 nt open reading frame
encoding 437 amino acids and a 309 nt 39 untranslated
region. No transit peptide could be identified from
the amino acid sequence and analysis with the PSORT
program showed high probability of a cytosolic localization. The 1608 bp nucleotide sequence of the mRNA
isolated for GSH-S (EMBL Nucleotide Sequence Database Acc. No. AJ302784) presents an open reading frame
between nucleotides 278 and 1510 that encodes a peptide
of 409 amino acids, with no transit peptide. Southern
blot analysis indicates that the gene encoding GSH-S
is present as a single copy. Screening of a BAC library
from the flint inbred F2 line with probes obtained in this
work confirmed this result. Although the analysis of
several clones of the c-ECS cDNA showed only one
sequence of mRNA, the screening of the BAC library
indicated that this gene is present in two copies. Northern
blot analysis showed that the expression of both enzymes
is higher in leaves than in roots but there are no results
to date on the intracellular regulation of the expression
of these genes.
Intracellular compartmentation of
glutathione metabolism
Intercompartmental variations (e.g. chloroplast versus
cytosol, apoplast versus cytosol) in glutathione concentration and redox state may be crucial in signalling.
By the end of the 1970s, it was known that spinach
chloroplasts contained high concentrations of glutathione
(Foyer and Halliwell, 1976) and that photoautotrophic
tobacco cells exported glutathione into the culture
medium much faster than did hetrotrophically grown
cells (Bergmann and Rennenberg, 1978). It was subsequently shown that GSH is translocated from source
leaves in the phloem (see references in Herschbach et al.,
2000). As discussed above, c-ECS and GSH-S activities
were shown to be located both inside and outside the
chloroplast (Klapheck et al., 1987; Hell and Bergmann,
1988, 1990) and the ability of photosynthetic cells to
synthesize GSH in chloroplastic and cytosolic compartments was confirmed by overexpression studies ( Noctor
et al., 1996, 1998; Creissen et al., 1999). Together, these
observations indicate that photosynthetic cells are likely
to be able to export chloroplastically-produced glutathione from the cell. Studies have been conducted of
35
S-GSH uptake into tobacco cells and bean leaf
protoplasts (Schneider et al., 1992; Jamaı̈ et al., 1996),
and a GS-conjugate transporter is known to operate at
the tonoplast (Foyer et al., 2001), but nothing is known
about glutathione transport across the chloroplast
envelope. The following sections present results that
have recently been obtained on compartmentation and
1289
transport, and these findings are discussed within the
context of published literature data in order to evaluate the role of different processes in the control of the
intracellular distribution of glutathione metabolism.
Distribution of glutathione metabolism between the
chloroplast and the rest of the cell
Because of the agronomic importance of wheat, the
compartmentation of glutathione metabolism was undertaken by isolating intact chloroplasts from wheat leaves.
Although highly intact wheat chloroplasts can be prepared by lysis of protoplasts prepared via enzymatic
digestion (Edwards et al., 1978), this method is of questionable suitability for the study of the distribution of
stress-linked components because the isolation procedure
itself affects AOS production and the antioxidant system
(Ishii, 1987; Papadakis et al., 2001). Therefore, the development of a more direct and rapid method for the
isolation of intact wheat chloroplasts was sought, using
the mechanical homogenization of leaf tissue. While
mechanical homogenization is easily applied to species
with soft leaf tissue (notably spinach and pea), the
literature contains very few reports of the successful use
of this technique in the isolation of wheat chloroplasts,
probably because fragments of fibrous bundle-sheath
strands are released during tissue homogenization and
cause rupture of the fragile chloroplasts. Nevertheless, by
using young leaves, it was possible to develop a method
producing an adequate yield (0.5–1 mg chlorophyll) of
predominantly intact chloroplasts (Table 2). This method
allowed chloroplasts to be liberated and separated from
the homogenate in under 5 min and to be purified in less
than 30 min. Given that all leaf antioxidative enzymes,
including chloroplastic isoforms, have thus far been
shown to be nuclear-encoded, this rapid isolation technique should avoid possible artefactual changes in
enzyme distribution. The chloroplasts were photosynthetically competent, as judged by their high rates of electron
transport (Table 2). Although unable to catalyse CO2dependent O2 evolution at appreciable rates, the chloroplasts were competent in O2 evolution when supplied
with 3-phosphoglyceric acid (Table 2). This indicates
activity of the phosphate translocator and at least two
enzymes of the Calvin cycle, as well as retention of
nucleotides ( NADP(H), adenylates). The chloroplast
volume (Table 2) was within the range of that measured
in other species (Heldt, 1980).
Expressed with respect to chlorophyll, NADPGAPDH activity in the wheat chloroplasts was similar
to that measurable in the unfractionated homogenate
(Table 3). The latency of this activity in the chloroplast
preparation was greater than 80%. While a large part of
GR activity was located in the chloroplasts, the proportion of DHAR associated with the organelle was rather
1290
Noctor et al.
Table 2. Characterization of chloroplasts isolated mechanically
from young wheat leaves
Data for chloroplasts from pea leaves, isolated by a similar protocol, are
shown for comparison. Values are means ] SD of 23 (wheat) and three
( pea) independent chloroplast preparations (single measurement of
chloroplast volume in pea). Leaves (7 d after sowing for wheat, 12 d
after sowing for pea) were ground in iso-osmotic buffer in a Polytron
homogenizer, filtered and chloroplasts were rapidly pelleted by
centrifugation. The pellet was resuspended in iso-osmotic buffer and
intact chloroplasts were purified by centrifugation for 5 min through a
40% percoll cushion. The final pellet was washed, then resuspended to
a chlorophyll concentration of 0.5–1 mg ml Y 1 and used to obtain the
data shown below, as well as those in Tables 1, 3 and 4, and Figs 4 –7.
The maximum rate of non-cyclic electron transport was measured in
osmotically shocked chloroplasts in the presence of 5 mM ferricyanide
and 10 mM NH4Cl at an irradiance of 1000 mmol m Y 2 s Y 1. The
chloroplast volume was measured in the dark using 14C-sorbitol and
3
H2O (according to Heldt, 1980). PGA-dependent and CO2-dependent
O2 evolution were measured at an irradiance of 600 mmol m Y 2 s Y 1 in
intact chloroplasts supplemented with 5 mM sodium iso-ascorbate,
0.3 mM KH2PO4, 1 mM PGA or 5 mM NaHCO3. All measurements
were carried out at 20 C; n.m., not measured.
Wheat
Maximum rate electron transport
(mmol O2 mg Y 1 chl h Y 1)
% Intactness
(ferricyanide exclusion)
Chloroplast volume
(ml mg Y 1 chl)
PGA-dependent O2 evolution
(mmol mg Y 1 chl h Y 1)
CO2-dependent O2 evolution
(mmol mg Y 1 chl h Y 1)
Pea
272 ] 64
229 ] 17
78.5 ] 7.4
98 ] 3.5
30.7 ] 6.5
21.9
68 ] 14
n.m.
< 20
53 ] 13
Table 3. Distribution of enzymes associated with glutathione
metabolism between the chloroplastic and extra-chloroplastic
compartments of young wheat leaves
NADP-dependent glyceraldehyde-3-phosphate dehydrogenase (NADPGAPDH) was measured, without prior activation, as PGA-dependent
NADPH oxidation in the presence of phosphoglycerate kinase
and ATP (Foyer and Halliwell, 1976). Glutathione reductase (GR)
and dehydroascorbate reductase (DHAR) were measured at 25 C
(by methods adapted from Foyer and Halliwell, 1976). Glutathione
synthetase (GSH-S) was measured under anaerobic conditions at 30 C
(by a method adapted from Hell and Bergmann, 1988), and very similar
to that described in Fig. 2 for c-ECS, except that Cys was replaced by
c-EC (1 mM) and Glu was replaced by Gly (2 mM). The reaction was
started by addition of Gly, and samples were taken at 5, 25, and 45 min
after addition. c-EC and GSH were separated according to the same
HPLC protocol as that used for separation of Cys and c-EC (Fig. 2).
Rates were obtained by subtracting the amount of GSH formed after 25
and 45 min from that present after 5 min. Values are the means ] SD of
three independent chloroplast preparations except for data for DHAR,
which are the means of two preparations. All rates are expressed in
mmol mg Y 1 chl h Y 1.
Enzyme
Purified
chloroplasts
Unfractionated
homogenate
% found in
chloroplast
NADP-GAPDH
GR
DHAR
GSH-S
78 ] 18
37.2 ] 1.0
70
0.07 ] 0.02
72 ] 21
66.6 ] 4.7
306
0.78 ] 0.15
108
56
23
9
less (Table 3). The fraction of GSH-S found in the
chloroplasts was low (Table 3). The use of chlorophyll to
calculate the distribution of enzymes within the photosynthetic cell is complicated by the existence of significant
numbers of non-photosynthetic cells within leaves. In
an analysis of wheat of the same age as that used here,
only 50% of the leaf cells were found to be mesophyll
cells, with only slightly higher values found in other
monocotyledonous and dicotyledonous species, including
spinach and pea (Jellings and Leech, 1982). Values for
the chloroplastic enzyme fraction based on chlorophyll
(Table 3) must therefore be treated as lower limits for
the chloroplast allocation within photosynthetic cells:
true values would be higher if a significant fraction of
enzyme is also present in non-photosynthetic cells.
Four thiols were detected in wheat leaves and
chloroplasts (Table 4). The glutathione homologue
c-Glu-Cys-Ser (hmGSH) was first described in 1992 and
reported to exist in numerous grass species (Klapheck
et al., 1992). It was not possible to detect the formation of
this compound in GSH-S assays where Gly was replaced
by Ser, consistent with this thiol being produced by
hydroxymethylation of the Gly residue of GSH. Contents
of all thiols on a chlorophyll basis were much lower in
chloroplasts, only 8% of the total leaf glutathione being
recovered in the purified chloroplasts (Table 4). Assuming that the vacuole occupies 80–90% of the cell volume,
and that glutathione concentration is low in this compartment (Wolf et al., 1996), a leaf content of 199 nmol
mg Y 1 chl (Table 4) yields a global concentration outside
the chloroplast of 1–2 mM (chlorophyll content of the
wheat leaves was around 1 mg g Y 1 FW ). This value is
in agreement with estimations of root cytosolic concentration using in situ imaging techniques (1.8– 4 mM:
Fricker and Meyer, 2001). The glutathione content of
the isolated chloroplasts (Table 4) can be converted
directly, using the measured chloroplast volume (Table 2),
to a concentration of around 0.5 mM. This would suggest
that the glutathione concentration in young wheat leaves
is lower in the chloroplast than in other compartments
such as the cytosol. Several observations speak against
Table 4. Chloroplasts from wheat leaves retain only a small
proportion of thiols present in the leaf
Thiols were extracted from chloroplasts and leaves by acid extraction
into 0.1 M HCl, 1 mM EDTA and analysed by reverse-phase HPLC
with fluorimetric detection of monobromobimane derivatives (by a
method modified from Noctor and Foyer, 1998). Contents are expressed
in nmol mg Y 1 chl (means ] SD of three independent leaf extracts or
chloroplast preparations). hmGSH, hydroxymethyl glutathione (c-GluCys-Ser).
Leaf extracts
Chloroplasts
% in Chloroplast
Cysteine
c-EC
hmGSH
GSH
30.1 ] 3.1
5.0 ] 0.3
17
8.0 ] 2.4
0.24 ] 0.03
3
110 ] 19
4.2 ] 1.0
4
199 ] 15
15.5 ] 1.2
8
this notion. First, a higher chloroplast glutathione
concentration (63–81 nmol mg Y 1 chl) was reported in
spinach, representing a mean concentration of 3.5 mM
(Foyer and Halliwell, 1976). Second, a higher proportion of glutathione was also found in chloroplasts isolated non-aqueously from barley (Smith et al., 1985).
Here, more than 50% of leaf glutathione was recovered
in the chloroplasts, although this method may lead to
overestimation due to adhesion of extrachloroplastic
material to the isolated chloroplasts. Using marker
enzymes to correct for this artefact, a value of 35% leaf
glutathione in pea chloroplasts isolated in non-aqueous
media was calculated (Klapheck et al., 1987). By contrast,
the same article reported that chloroplasts prepared in
aqueous media from pea protoplasts retained only 5% of
glutathione present in the parent protoplasts (Klapheck
et al. 1987). When isolated in aqueous media, pea chloroplasts had glutathione contents of 7–22 nmol mg Y 1 chl,
values very similar to those we found in wheat
(cf. Table 3). Moreover, another study in pea found only
10% of the leaf glutathione in percoll-purified chloroplasts (Bielawski and Joy, 1986). It was concluded that
pea chloroplasts lose glutathione during extraction
in aqueous media, probably due to leakiness to small
molecules during the isolation (Klapheck et al., 1987).
Another possibility is the operation of transporters
(see below). Whatever the processes responsible for low
contents following aqueous isolation, it seems clear that
chloroplasts from wheat, like those from pea, lose
glutathione and other thiols during this method of
extraction while retention of glutathione is higher in
spinach chloroplasts. Interestingly, it was reported
that, expressed on a chlorophyll basis, the glutathione
content of pea protoplasts was similar to that of
pea leaves (Klapheck et al., 1987). By contrast, mesophyll protoplasts have been purified from wheat
(authors’ unpublished results) that have high photosynthetic activity (100–170 mmol O2 mg Y 1 chl h Y 1 in the
presence of NaHCO3) but that contain only approximately 30% of the initial leaf glutathione content (c. 60
and 200 nmol mg Y 1 chl in protoplasts and leaves,
respectively).
Table 5 presents a summary of the principal literature
data on the chloroplast complement of glutathione,
together with values for the four enzymes that were
measured in wheat. In the leaves of dicotyledonous
species, c-ECS and GSH-S have been found to be more
or less equally divided betwen chloroplastic and nonchloroplastic compartments, with a somewhat higher
proportion of c-ECS found in the chloroplast (Table 5).
The effects of protein concentration on c-ECS activity
(Fig. 2) prevented estimating the distribution of this
enzyme in wheat leaves, but it seems clear that, whatever the precise distribution, a higher proportion of this
enzyme is found in the wheat chloroplast than GSH-S.
1291
Table 5. Literature data for the fraction of tissue glutathione and
associated enzymes found in plastids
Enzyme
Species and tissue
% in plastid
c-ECS
Pea leaves
Spinach leaves
Maize roots
Cultured tobacco cells
Pea leaves
Spinach leaves
Maize roots
Pea leaves
Spinach leaves
Spinach leaves
Pea leaves
Pea leaves
Barley leaves (chloroplasts
isolated in non-aqueous media)
72b
61b
46d
24a
48b, 47–69c
58b
8d
52–75e, 77h, 77i
80g, 67h
9g, 28h
65i
10e, 5c, 35c
50–65 f
GSH-S
GR
DHAR
Glutathione
a
Hell and Bergmann, 1988.
Hell and Bergmann, 1990.
c
Klapheck et al., 1987.
d
Rüegsegger and Brunold, 1993.
e
Bielawski and Joy, 1986.
f
Smith et al., 1985.
g
Calculated from Foyer and Halliwell, 1976.
h
Anderson et al., 1983.
i
Gillham and Dodge, 1986.
j
Edwards et al., 1990.
b
In nmol chloroplast protein Y 1 min Y 1, maximal activities
were 2.3 (c-ECS: Fig. 2) and 0.23 (GSH-S). This is
surprising considering that overexpression studies
strongly suggest that the first enzyme exercizes the
major limitation on synthesis of GSH in both cytosolic
and chloroplastic compartments (Strohm et al., 1995;
Noctor et al., 1996, 1998). However, studies of spinach
chloroplasts, and of barley and pea chloroplasts isolated
non-aqueously, indicate a chloroplastic GSH concentration close to 5 mM (Foyer and Halliwell, 1976; Smith
et al., 1984; Klapheck et al., 1987) which would be
sufficient to inhibit the chloroplastic c-ECS by about 90%
(Fig. 3) at 10–20 mM Glu, a likely chloroplastic concentration of this amino acid (Winter et al., 1994). It is
perhaps worth noting that analysis of maize roots also
found only a small fraction of GSH-S in the plastid
(Table 5). The distribution of these enzymes could be
dependent on developmental stage. Even though the
wheat leaves used here were photosynthetically competent, they were nevertheless young. Literature data suggest, perhaps, that in more mature leaves, GSH-S is more
strongly associated with the chloroplast (Table 5).
The data presented in Table 2 for GR are in agreement
with literature studies (Table 5). In leaves, the bulk of GR
activity is found in the chloroplast whereas root plastids
may contain a lower proportion of the total cellular
activity (Table 5). About 20% of the pea leaf activity was
associated with the cytosol (Edwards et al., 1990). Lower
GR activities have been reported in isolated mitochondria
and peroxisomes (Edwards et al., 1990; Jimenez et al.,
1997). Of the four enzymes discussed here, GR is by far
1292
Noctor et al.
the best characterized at the gene level. The first gene
sequence encoding plant GR was isolated from pea,
shown to encode a product with an N-terminal sequence
characteristic of chloroplast-targeting sequences (Creissen
et al., 1992), and later the product was found to be
targeted to both chloroplasts and mitochondria (Creissen
et al., 1995). Subsequently, cDNAs for other isoforms
have been isolated from various species. Multiple activity
bands have been identified in protein extracts from pea
and spinach (Edwards et al., 1990; Foyer et al., 1991).
Less is known about the compartmentation of DHAR.
The data in wheat (Table 3) give a value intermediate
between those found in spinach and pea (Table 5).
Because various enzymes can catalyse GSH-dependent
reduction of dehydroascorbate as a ‘secondary’ reaction,
the presence of a specific DHAR activity in chloroplasts has been the subject of controversy (Foyer and
Mullineaux, 1998). Very recently in spinach, however, a
chloroplast DHAR has been purified and a corresponding gene cloned and sequenced (Shimaoka et al., 2000).
The authors reported a stromal DHAR activity of
34 mmol mg Y 1 chl h Y 1 (Shimaoka et al., 2000). Since
DHAR activities from leaf tissue are typically
100– 400 mmol mg Y 1 chl h Y 1, this would also represent
a fairly small proportion associated with the chloroplast, although part of the high extrachloroplastic
activity may be attributable to proteins other than a
specific DHAR. It is interesting that GRuDHAR ratios
appear to be markedly higher in the chloroplast than
outside this organelle: on the basis of differential effects
on the ascorbate and glutathione redox states observed
in transgenic and mutant plants, it was suggested that
DHAR activity in the chloroplast may be too low to
couple the ascorbate and glutathione pools effectively
(Noctor et al., 2000). In a recent theoretical article, the
first attempt to model flux through the chloroplast
Mehler-peroxidase and ascorbate–glutathione cycles is
reported (Polle, 2001). It was concluded that chloroplastic
DHAR is likely to be unimportant in chloroplast redox
cycling, but that the chemical reduction of DHA by GSH
would be sufficiently fast to allow an effective ascorbate–
glutathione cycle (Polle, 2001). It is nevertheless clear that
the redox states of the leaf ascorbate and glutathione
pools can vary independently: decreased expression of
the chloroplastic protein 2-cys peroxiredoxin led to a
more oxidized ascorbate pool without effect on the highly
reduced glutathione pool (Baier et al., 2000). Both
thermodynamic and kinetic considerations predict that
glutathione should be oxidized before ascorbate (Noctor
et al., 2000; Polle, 2001), and this is indeed observed in
plants deficient in CAT (see below). The factors that may
allow DHA to accumulate in the presence of abundant
GSH have been discussed, and include microcompartmentation within the chloroplast (Noctor et al., 2000;
Polle, 2001). Polle’s model is an excellent first step
towards the comprehensive evaluation of oxidant processing and antioxidant cycling within the chloroplast,
and presents or reinforces several important conclusions
concerning the importance of non-enzymatic reduction
and the likely location of ‘rate-limitations’ (Polle, 2001).
Such approaches are likely to be very useful in understanding the response of chloroplast-derived stress to
leaf physiology. Nevertheless it is noted that the common
assumption that the chloroplast is the predominant site
of AOS production in leaf cells may not be valid under
many conditions (Noctor et al., 2002), and that it cannot
be excluded that the chloroplast antioxidative system
may sometimes be subject to oxidative loads of both
chloroplastic and extra-chloroplastic origin.
Transport of glutathione across the chloroplast envelope
It has previously been reported that incubation of wheat
chloroplasts with 35S-labelled GSH at 1 and 100 mM
resulted in time-dependent uptake that was linear for at
least 15 and 8–10 min, respectively (Noctor et al., 2000).
To characterize the uptake process, rates were measured
over the first 5 min following addition of 35S-GSH, within
which time uptake was linear at all concentrations
between 1 mM and 1 mM (Fig. 4). Regression analysis
Fig. 4. Time-courses for uptake of different concentrations of 35S-GSH
into intact wheat chloroplasts. 35S-GSH was obtained from NEN
biolabs (Boston, USA) and checked for purity by HPLC separation of
monobromobimane-labelled thiols followed by scintillation counting
of collected fractions. Contamination with cysteine and c-EC was 0.6%
and 1.8%, respectively. Chloroplasts were incubated at a chlorophyll
concentration of 0.1 mg chl ml Y 1 in isotonic buffer ( pH 7.6) at 25 C in
the dark. Uptake was initiated by addition of GSH to 1 m Ci ml Y 1 35SGSH and chemical concentration as indicated. At the times shown,
aliquots of 0.1 ml were withdrawn and intact chloroplasts were rapidly
pelleted by centrifugation for 5 s through 0.1 ml silicone oil (AR200,
Wacker Chemie GmbH, Munich, Germany) into 0.02 ml 1 M HClO4.
The tubes were cut just above the oil–acid interface with a razor blade
and radioactivity in the acid pellet was determined by scintillation
counting. The chloroplast content of glutathione was determined by the
measurement of 14C-sorbitol-permeable and 3H2O-permeable spaces
(according to Heldt, 1980). Note the different scales on the y-axes of
different panels.
was used to calculate rates: fitted lines gave a positive
intercept on the y-axis (Fig. 4), probably indicating a
rapid binding to the external surface of the chloroplast
followed by a constant rate of uptake. At low concentrations (1–50 mM) the time-dependent uptake results in an
accumulation of glutathione to a calculated chloroplast
concentration of up to 5-fold the external concentration. Given that these external concentrations represent
values at least 10-fold lower than the internal chloroplast concentration before addition of labelled GSH
(c. 0.5 mM: see above), the data suggest active uptake
of external glutathione. This conclusion is supported by
the concentration dependence of GSH uptake (Fig. 5).
Uptake was linear up to 20–30 mM GSH then showed
saturation at around 100–200 mM GSH followed by a
further increase in rate up to a concentration of 1 mM
(Fig. 5). The results suggest that at least two systems
are able to take up GSH across the chloroplast envelope,
one showing half-saturation at around 30–50 mM
GSH with a maximal capacity of approximately
0.6 – 0.8 nmol mg Y 1 chl min Y 1 and a second with lower
affinity and higher capacity. Transport at 50 mM GSH
was not affected by added ATP or by light (data not
shown). No difference in uptake was observed if the
possible oxidation of GSH was countered by the presence
of NADPH and yeast GR in the external medium.
Although direct uptake of the disulphide form has not
been examined, the presence of GSSG at 10 and 500 mM
significantly inhibited uptake of GSH at 10, 50 and
500 mM. At a constant GSH concentration of 50 mM,
inhibition by GSSG was half-maximal at approximately
0.4 mM (Fig. 6). These data suggest that the two forms
of glutathione can be transported by common systems,
although GSH appears to be preferred. GSSG was also
reported to inhibit GSH uptake by bean protoplasts
(Jamaı̈ et al., 1996), but not by tobacco cells (Schneider
et al., 1992). It is conceivable that the weak effect of
GSSG in wheat chloroplasts (Fig. 6) could be due to an
effect on 35S-GSH concentration rather than an effect
on the transport process itself. However, mixtures of
GSSG and GSH are relatively stable at neutral pH so
an effect of GSSG on the chemical GSH concentration
is unlikely given the brevity of each incubation. It is
possible, nonetheless, that thiol-disulphide exchange
reactions result in conversion of 35S-GSH to 35S-GSSG,
particularly when GSSG is in excess of GSH. Here, it
is worth noting that Schneider reported no effect of
0.25 mM GSSG on the uptake of 35S-GSH (50 mM) into
tobacco cells during a 2 h incubation at pH 6.0 (Schneider
et al., 1992). Although this study’s experiments were
conducted at pH 7.6, which is likely to be more conducive to thiol-disulphide exchange, all solutions were prepared immediately before assay and the incubation period
used (5 min) was much shorter than those employed
previously in studies of glutathione uptake across the
1293
Fig. 5. Concentration dependence for uptake of 35S-GSH by intact
wheat chloroplasts. Rates were calculated by linear regression analysis of
curves as shown in Fig. 4. Apart from GSH concentration \ 1.5 mM
(single measurement), all values show the means ] SE of rates
determined on between two and eight independent chloroplast preparations. The plot uses 52 separate measurements of rates in 12 independent
chloroplast preparations.
Fig. 6. Inhibition of GSH uptake by GSSG. GSH was held constant at
50 mM. Methods as in Figs 4 and 5.
plasmalemma. Nevertheless, it remains unclear whether
the effect of GSSG is mediated at the level of transport.
The capacity of the high-affinity wheat chloroplast
glutathione uptake system was 40–60 nmol mg Y 1 chl h Y 1
1294
Noctor et al.
while, at physiological GSH concentrations (1 mM), the
uptake rate was about 200 nmol mg Y 1 chl h Y 1 (Fig. 5).
By applying a modified but similar method, it was not
possible to detect significant uptake into wheat mesophyll
protoplasts within 60 min, at either pH 7.6 or pH 5.5.
This contrasts with the literature report of significant
GSH uptake into broad bean protoplasts, where transport was shown to be due to a single saturable phase with
Km 0.4 mM and Vmax of 2.1 nmol 107 protoplasts min Y 1
(Jamaı̈ et al. 1996). This difference may be attributable to
differences in methodology or to inactivation of the wheat
plasmalemma translocator during protoplast isolation.
It is worth comparing maximum GSH transport rates
across the plasmalemma with those measured in wheat
chloroplasts. A mean chl content per wheat protoplast of
11.8 ] 2.0 pg (n \ five independent protoplast preparations) was measured. Assuming a similar chl content, a
rate of 2.1 nmol Z 107 broad bean protoplasts min Y 1
converts to around 1 mmol mg Y 1 chl h Y 1, though this
capacity is unlikely to be reached at physiological
extracellular GSH concentrations. In photoheterotrophic
tobacco cells, as in wheat chloroplasts, two phases were
identified in kinetic characterization of GSH uptake
(Schneider et al., 1992). A high-affinity system displayed
a Km of 18 mM and a capacity of 19–20 nmol GSH g Y 1
dry weight min Y 1, while at higher concentrations a
second phase was observed with Km 780 mM and a
capacity of c. 170 nmol GSH g Y 1 dry weight min Y 1
(Schneider et al., 1992). The physiological significance
of the second system is unclear. Given a chl content in
the region of 10–20 mg g Y 1 dry weight, high-affinity
transport by the tobacco cells would proceed at about 60 –
120 nmol mg chl Y 1 h Y 1. The capacity of wheat chloroplast transport is therefore intermediate between those
previously measured in photoheterotrophic tobacco cells
and broad bean protoplasts.
The bean protoplast plasmalemma transports GSSG
at higher rates than GSH (Jamaı̈ et al., 1996), consistent
with a predominant physiological role in the recovery of
glutathione oxidized in the apoplast, where GR activity is
low or nil. Further work is required to identify possible
chloroplast envelope transporters which might preferentially transport GSSG. It remains unclear whether
GSSG and GSH are transported by the same system at
the plasmalemma. The two species had distinct kinetics,
GSH showing a single saturable phase with Km 0.4 mM
whereas GSSG showed two saturable phases (Jamaı̈ et al.,
1996). Protoplast transport of the two species were
similarly dependent on pH and both resulted in decreased
acidification of the extracellular medium in leaf pieces
(Jamaı̈ et al., 1996).
A high-affinity glutathione transporter has recently
been cloned from yeast, with a Km value close to that of
the high-affinity system characterized in tobacco cells
and wheat chloroplasts (54 mM: Bourbouloux et al.,
2000). Negligible inhibition was observed with other
peptides or amino acids, but glutathione conjugates and
GSSG both competed significantly with GSH. A database
search of amino acid sequences identified homologues
from other organisms, including five from Arabidopsis,
which had 38–51% identity with the cloned yeast gene
(Bourbouloux et al., 2000).
Although glutathione concentrations in the vacuole are
thought to be low, it is clear that the compound enters via
transport of conjugates. Transport from the cytosol to the
vacuole occurs via a Mg-ATP glutathione-S-conjugate
transporter which is up-regulated along with GSTs upon
exposure to xenobiotics (Martinoia et al., 1993; Li et al.,
1995). The glutathione S-conjugates formed with anthocyanin and medicarpin are also transported into the
vacuole by a specific glutathione S-conjugate transporter
(Li et al., 1995), where they are further metabolized
(Marrs, 1996; for further discussion of vacuolar uptake,
see Foyer et al., 2001).
A key question concerning glutathione transporters
is whether they catalyse net transport. Many plastid
envelope translocators involved in primary C and N
metabolism generally catalyse strict exchange reactions
(e.g. phosphate translocator, dicarboxylic acid transporter, glutamate transporter, adenylate translocator).
An obvious candidate for exchange with glutathione is
not apparent, and further work is required to resolve this
question. If an exchange mechanism is not involved, a
unidirectional transport of glutathione would presumably require energy input. Evidence was obtained that
uptake of both GSH and GSSG into bean protoplasts
was driven by the proton gradient across the plasmalemma (Jamaı̈ et al., 1996). On the other hand, the rate
of GSH uptake into photoheterotrophic tobacco cells by
the high-affinity system decreased as the pH was lowered
from 7.0 to 5.0 (Schneider et al., 1992). The pH gradient
across the chloroplast envelope is smaller than that across
the plasmalemma and is very light-dependent (Werdan
et al., 1975). The mechanism of chloroplast envelope
transport of glutathione requires further investigation.
Another interesting question is whether specificity to
glutathione, observed in plasmalemma transport (Jamaı̈
et al., 1996; Bourbouloux et al., 2000), is conferred by
the Glu-Cys c-carboxy peptide bond, or whether the
transporter recognizes the thiol group. The work of
Schneider et al., which reported inhibition by pretreatment of the cells with the alkylating reagent iodoacetate but not by the presence of GSSG during the
uptake assay, suggests that the transport may involve
thiol-binding (Schneider et al., 1992). On the other hand,
the results of Jamaı̈ et al. suggest that the characteristic
N-terminal peptide link may be crucial (Jamaı̈ et al.,
1996). It is interesting to note, however, that the cloned
yeast transporter showed little activity with c-EC
(Bourbouloux et al., 2000).
Glutathione homeostasis: the relationship
between concentration and redox state
It is an interesting question whether many plant processes
are subject to strict homeostasis in the sense in which this
term is applied to mammalian systems. Plant metabolism
is generally less insulated from environmental variation
and, as a result, has evolved considerable flexibility and
redundancy. Two independent properties of any given
pool of glutathione are obvious: its concentration and
its redox state. The first, although subject to regulation
at multiple levels (notably, as far as homeostasis is concerned, by end-product feedback inhibition), can vary
considerably. Leaf glutathione concentrations fluctuate
seasonally, diurnally, and are influenced by plant nutrition, particularly sulphur availability. Earlier evidence
that glutathione acts homeostatically in sulphur metabolism, being synthesized primarily in the leaves then
translocated in the phloem to regulate sulphate uptake at
root level, has been succeeded by a more complex picture
in which the phloem sulphate to glutathione ratio may be
key (Herschbach et al., 2000). The second property of
the glutathione pool, its redox state, is perhaps more
likely to be the subject of strict homeostatic control, as
part of the maintenance of general cellular redox poise. In
leaves at least, the glutathione redox state is remarkably
constant. Only under fairly sustained stress conditions
does the leaf glutathione pool fall much below 90%
reduced. The centrality of a highly reduced glutathione
pool to photosynthetic metabolism may be such that
departures from the highly reduced state are sensed by the
cell and acclimatory changes in gene expression initiated.
It is important to point out that redox state is a term
used widely, though sometimes without precise and correct meaning, in the glutathione research field (Schafer
and Buettner, 2001). The redox potential of a given redox
couple, existing in two forms, Ared and Aox, is given by the
Nernst equation.
E ¼ Em ððRT=nF ÞÞlnð½Ared =½Aox Þ
where E, redox potential of A; Em, midpoint potential of
A (where wAoxx \ wAredx); R, gas constant; T, temperature;
N, number of electrons involved in the interconversion
of Aox and Ared; F, Faraday constant.
It is clear from this equation that changes in
glutathione redox potential do not occur if GSHuGSSG
remains constant, even if the total concentration of the
pool changes dramatically. Hence, the total glutathione
concentration and its redox state may be independent
parameters, from a chemical point of view. However,
signalling initiated by changes in the redox state may
lead to up-regulation of glutathione synthesis and, hence,
increases in the total concentration. A likely redox potential for the chloroplastic glutathione redox couple is
Y 0.23 V (Foyer and Noctor, 2000). The glutathione
1295
redox potential of animal cells has been estimated to
vary from about Y 0.24 V in actively dividing cells to
approximately Y 0.17 V in cells undergoing apoptosis
(Schafer and Buettner, 2001).
Maintenance of homeostasis in the face of all the
metabolic demands placed on the glutathione pool
involves a complex interplay between synthesis, degradation, transport, storage, oxidation–reduction, further
metabolism and catabolism as plants respond to environmental, developmental and nutritional cues. Redox
cycling is much faster than synthesis, transport or
degradation. However, the antioxidant systems ensure
that any changes in glutathione redox state are relatively
slow. Although it is difficult to know the rate of redox
turnover of glutathione in vivo (though see Polle, 2001,
for a thorough analysis of flux through the chloroplast
antioxidative system), plants deficient in CAT display
probably the best documented and most striking perturbation of the glutathione pool. Low activities of the
major leaf peroxisomal form of this enzyme mean that
the copious amounts of H2O2 generated in C3 plants in
photorespiration must be metabolized by an alternative
route. This imposes a marked increase in the oxidative
load on the photosynthetic cell which, ultimately, results
in cell death and necrosis (Smith et al., 1984; Willekens
et al., 1997). For a limited period of a few days, however, barley CAT mutants can cope with photorespiratory
H2O2 without showing deleterious effects on either
photosynthesis or phenotype (Smith et al., 1984; Noctor
et al., 2000). Their ability to do so presumably reflects
enhanced engagement of other pathways of H2O2
detoxification, notably the ascorbate–glutathione cycle,
as evidenced by the sustained oxidation of glutathione
accompanied by increases in the total glutathione pool. In
the barley mutant, the net accumulation of GSSG 4 d
after transfer to air was around 0.7 mmol g Y 1 FW (Smith
et al., 1984). Very similar effects are observed in
transformed tobacco deficient in CAT (Willekens et al.,
1997). At least in barley, H2O2 does not accumulate to
any great extent within this time (Noctor et al., 2001).
Since the mutant has negligible leaf CAT activity and
rates of H2O2 generation via photorespiration can be
estimated relatively easily, a rough estimate can be
made of the relationship between redox cycling and
net oxidation of the glutathione pool. In a recent study
of this mutant (Noctor et al., 2002), results very similar to those of Smith et al. were obtained (Smith
et al., 1984). These experiments were carried out at
250 mmol quanta m Y 2 s Y 1, at which a typical rate of
ribulose-1,5-bisphosphate oxygenation in barley leaves
is around 180 mmol g Y 1 FW h Y 1 (Noctor et al., 2002:
for comparison, typical whole leaf GR capacity
measured under standard conditions is about
70 mmol g Y 1 FW h Y 1). At this rate, the photorespiratory
process generates approximately 10 mmol H2O2 g Y 1 FW
1296
Noctor et al.
in 4 d with a 14 h light period. The proportion of
H2O2 detoxification that involves redox turnover of the
glutathione pool is difficult to estimate. Polle concluded
that in the chloroplast this value was probably low and
that oxidation of glutathione by chemical reaction with
superoxide was equally if not more important than GSH
oxidation by DHA (Polle, 2001). Outside the chloroplast,
however, in CAT-deficient plants, glutathione may be
much more closely involved in H2O2 detoxification: (1)
in the illuminated leaves of C3 plants, under many
conditions, the major source of AOS production outside
the chloroplast is the direct production of H2O2 by
glycollate oxidase activity; (2) although monodehydroascorbate (MDHA) reductase activity is found outside
the chloroplast, ferredoxin-dependent regeneration of
ascorbate from MDHA is not possible; (3) in most
cases, as discussed above, extrachloroplastic DHAR
activities are higher than those within the chloroplast;
(4) extrachloroplastic GPXuGST activities may make a
more substantial contribution to H2O2 processing outside
the chloroplast. Even if it assumed that, when CAT
activity is negligible, only 1–10% of the H2O2 generated in
photorespiration drives oxidative turnover of glutathione
(either directly or via the ascorbate–glutathione couple),
the net accumulation of GSSG is still only around
0.07– 0.7% of the rate of redox turnover of the glutathione
pool. This figure will be lower if a higher proportion
of H2O2 detoxification involves engagement of the
glutathione pool. It is evident that accumulation of
GSSG can result from small imbalances in the rates
of oxidation and reduction of glutathione. Although
rates of synthesis and transport are probably around
two orders of magnitude slower than redox cycling
(Fig. 7), they are likely to be much closer to the imbalance
between reduction and oxidation, and could therefore
exert some influence on glutathione redox state in a given
compartment.
The data of Smith et al. suggest that the changes in
glutathione concentration and redox observed in the
barley mutant occur both within and outside the chloroplast (Smith et al., 1985). Generation of H2O2 within
the peroxisome is therefore capable of influencing the
chloroplastic glutathione pool. Whether the changes in
chloroplastic glutathione redox state are due to movement of H2O2 into the chloroplast or transport of GSSG
from the cytosol is unclear. The increase in the glutathione pool presumably results from up-regulation of
synthesis, perhaps through effects on translation of
c-ECS anduor GSH-S, as discussed above. It is worth
considering the factors that determine the intracellular
distribution of glutathione (Fig. 7). Although the capacity
of chloroplast transport is in the same range as the
maximum extractable activities of the enzymes that
catalyse glutathione synthesis, these data suggest that
Fig. 7. Control of the intracellular concentration of glutathione in leaf cells. For synthesis, transport and redox cycling, an estimate of likely rates
in vivo is indicated. GSH, reduced glutathione; GSSG, glutathione disulphide.
the transporter would operate much closer to its maximum capacity than the enzymes under in vivo conditions.
Under most conditions, the enzymes probably work at
considerably less than half maximal capacity because
of kinetically limiting substrate concentrations and competitive inhibition of c-ECS by glutathione. Increased
glutathione in one compartment will (a) counteract further glutathione synthesis through feedback inhibition
and (b) favour transport of glutathione out of that compartment, if net transport does occur. These processes
would tend to equalize glutathione concentrations between
chloroplast and cytosol, and compartment-specific
increases would require sustained differential increases in
the expression of enzyme isoforms. The extent to which
compartment-specific changes occur is unclear, but there
are several cytosolic processes (e.g. phytochelatin synthesis, GST activity) for which an increased synthesis
of chloroplastic glutathione would seem inappropriate.
However, it may well be that chloroplastic synthesis contributes to the supply of glutathione to these processes via
transport across the envelope. As noted above, emerging
evidence points to the existence of glutathione transporters with similar kinetic properties on different
membranes of the leaf cell. Perhaps the physiological
significance of these transporters is to minimize perturbations of glutathione homeostasis that would otherwise
result from differential demands in distinct compartments.
Catabolism of glutathione could also impact on
glutathione homeostasis. Three routes of breakdown,
involving GSH, GSSG and glutathione-S-conjugates, are
possible and each pathway may fulfil an essentially
different function. Catabolic destruction of GSSG may
serve as a detoxification process. GSSG is involved in
thiolation reactions forming mixed disulphides with proteins in conditions of oxidative stress. Since this process
inactivates many biosynthetic enzymes the presence of
a large GSSG pool is not compatible with many metabolic reactions; catabolism of GSSG would essentially
return the system to pre-stress homeostasis. Catabolism
of GSH, on the other hand, largely concerns the remobilization of cysteine, for example, during seed storage
protein synthesis or during periods of sulphur deprivation. This requires the successive breakage of the two
peptide bonds.
Glutathione catabolism is well characterized in
animals, failure of this process resulting in death (Meister,
1988). Transpeptidases, which catalyse the reversible
hydrolysis of the N-terminal peptide bond, initiate catabolism by removing the c-linked Glu from GSH, GSSG,
glutathione conjugates, and other peptides. The Glu
moiety is either hydrolysed or donated to an amino
acid acceptor or even to another GSH molecule. The
second step in catabolism is less well characterized. The
Cys–Gly bond is not unique to the glutathione tripeptide
and several enzymes, including aminopeptidase M and
1297
Cys–Gly dipeptidase, are able to hydrolyse the bond
(Meister, 1988). The transpeptidases are part of the
c-glutamyl cycle and as such are involved in amino acid
transport in some tissues ( Meister, 1988). The c-glutamyl
moiety is metabolized by a c-glutamylcyclotransferase
to oxo-proline which is subsequently converted to
glutamate by oxo-prolinase. Homologous activities are
also present in plants (Rennenberg et al., 1981; Steinkamp
et al., 1987; Steinkamp and Rennenberg, 1984). In addition, however, a carboxypeptidase exists which is able to
remove Gly as the first step of degradation, leaving c-EC
(Steinkamp and Rennenberg, 1985). These enzymes are
cytosolic but, more recently, a vacuolar carboxypeptidase
has been identified that cleaves the Gly moiety from
glutathione-S-conjugates (Wolf et al., 1996): it is thus
possible that cleavage of conjugated glutathione in the
vacuole may be a major route of catabolism in certain
conditions. Moreover, since GSSG can be considered
as a glutathione-S-glutathione conjugate, transport of
GSSG by the vacuolar conjugate transporter may play a
role in removing this species from the cytosol. The failure
to detect significant accumulation of glutathione-Sconjugates in vacuoles suggests that they are rapidly
catabolized in this compartment (Marrs, 1996).
Glutathione and signalling
The GSHuGSSG couple is well suited to the role of redox
sensor, indicative of the general cellular thiol-disulphide
redox balance, and producing profound effects on metabolism and gene expression. Regulation of gene expression by GSH and GSSG may be specific, i.e. these
compounds may be irreplaceable by other redox components. Alternatively, reported effects may reflect general
changes in the cellular redox state, which are known
to regulate gene expression in both prokaryotes and
eukaryotes.
The glutathione pool is an important redox component
in plant cells. Changes in intracellular glutathione status
may, therefore, be expected to have important consequences for the cell, through modification of the metabolic functions associated with glutathione-regulated
genes. In animal cells redox regulation of the transcription factor NFkB involves glutathione. This regulation is
important for T cell function since glutathione augments the activity of T cell lymphocytes (Suthanthiran
et al., 1990). Application of exogenous glutathione can
elicit changes in the transcription of genes encoding
cytosolic Cu,Zn superoxide dismutase and GR in tobacco
and pine (Hérouart et al., 1993; Wingsle and Karpinski,
1996) and 2-cys peroxiredoxins in Arabidopsis (Baier
and Dietz, 1997). Glutathione-inducible hypersensitive
elements have been identified in the proximal region
of the chalcone synthase (CHS) promoter (Dron et al.,
1988).
1298
Noctor et al.
Glutathione has been implicated in defence reactions
against biotic stresses. Marked changes in the glutathione
pool, such as those shown in Fig. 8, occur during the
hypersensitive response to pathogen attack. In barley
resistant to powdery mildew a transient decrease in the
leaf GSHuGSSG ratio, linked to H2O2 accumulation
around the mesophyll cells immediately below the
attacked epidermal cell, precedes the increase in the
total leaf glutathione pool (Fig. 8). Although pathogeninduced increases in the intracellular concentration of
glutathione and GSH-dependent induction of phenylalanine ammonia lyase and CHS have been demonstrated ( Wingate et al., 1988; Vanacker et al., 2000), it is
unlikely that GSH is the primary signal responsible
for the increase in phytoalexins following pathogen
attack. Using an artificial precursor of glutathione
Fig. 8. Induction of H2O2 and glutathione during the hypersensitive response in the leaves of resistant barley, in response to attack by the fungus
Blumeria graminis. The time-course of H2O2 accumulation in the attacked Alg-R leaf epidermal cells and in the underlying mesophyll was measured as
the appearance of brown coloration due to staining with diaminobenzadine. Cell death in epidermal cells was estimated by the accumulation of
autofluorogenic phenolic compounds. Specimens were stained post-fixation with aniline blue to show fungal structures. Micrographs were obtained
using transmitted white light (14.00–18.00 h) or incident blue-violet light (20.00–24.00 h). H2O2 was first detected 14 h after inoculation in mesophyll
cells underlying the attacked epidermal cell, at the same time as marked changes in the leaf glutathione pool were observed. A transient decrease in the
GSHuGSSG ratio preceded an increase in total leaf glutathione. These were specific to glutathione, the total leaf ascorbate content and AscuDHA ratio
remaining constant throughout pathogen attack. HR, hypersensitive response. (For other details see Vanacker et al., 2000).
biosynthesis, L-oxothiazolidine-4-carboxylate, to increase
intracellular thiol concentrations it was shown that
enhanced intracellular GSH concentrations alone did
not induce phytoalexin synthesis (Edwards et al., 1991).
It was concluded that changes in the intracellular glutathione concentration in response to pathogen attack
were too slow to be consistent with the initiation of the
elicitation response. In interactions between powdery
mildew and oat or barley, however, induction of glutathione precedes maximal induction of transcripts for
phenylpropanoid metabolism (Zhang et al., 1997;
Vanacker et al., 2000). There are differences in the signal
transduction pathways for elicitation of CHS transcription by fungal elicitor and glutathione (Choudhary et al.,
1990), suggesting that increases in glutathione are not
primarily responsible for the elicitation of the defence
response. A role for glutathione in subsequent parts of the
signal transduction pathway is nevertheless possible.
H2O2-mediated orchestration of gene expression may
be central to the ability of plants to elicit antioxidative
defences in response to abiotic and biotic threats and
hence develop pre-emptive cross-tolerance. Similarities
between the oxidative stress caused by ozone fumigation
and pathogen-induced responses suggest common elements in signal transduction routes involving SA, JA and
ethylene (Rao and Davis, 1999). Transformed tobacco
plants, deficient in the H2O2-scavenging enzyme CAT,
also show symptoms that are linked to the activation of
pathways involved in apoptosis, such as the induction of
pathogenesis-related (PR) proteins (Chamnongpol et al.,
1996). Hydrogen peroxide-mediated induction of glutathione has been demonstrated in different systems
(May and Leaver, 1993). It is apparent that the symptoms
which develop in CAT-deficient plants subjected to
prolonged exposure to photorespiratory conditions do
not simply result from chemical damage. Rather, they
resemble a precocious senescence, a regulated shutdown
of leaf cell function that involves components common
to signal transduction in response to pathogen attack
(Chamnongpol et al., 1996; Takahashi et al., 1997). A
central player is the phenolic molecule SA, but disruption of glutathione homeostasis may also be a vital piece
of the signalling jigsaw (Fig. 8).
AOS modulate nitric oxide signalling in the hypersensitive response, leading to cell death in the region
close to the pathogen attack (Delledonne et al., 1998)
and systemic acquired resistance (SAR) in surrounding
tissues. SAR involves the pre-emptive deployment of
gene expression to modify cell metabolism to cope with
future attacks. Increases in the SA concentration are
observed at the site of infection, and to a lesser extent at
remote sites. H2O2 and SA (and perhaps also glutathione)
are potential systemic messengers carrying information
concerning attack to unchallenged plant tissues. Of
the complex array of antioxidants found in plant cells,
1299
glutathione alone shows strong induction and rapid
accumulation in response to pathogen attack (Fig. 8;
Edwards et al., 1991; Vanacker et al., 2000). The effect on
biosynthesis is specific to glutathione and not a general
effect on the synthesis of low molecular weight antioxidants. It appears to be a universal response in plants
faced by pathogen attack or environmental stress, where
the antioxidant defences are temporarily overwhelmed by
an oxidative burst or by the accumulation of AOS as a
result of impaired metabolism.
The signalling mechanisms involved in induction of
GSH biosynthesis during pathogen attack are unknown.
As noted earlier, H2O2 increases tissue glutathione
contents, whereas JA increases the transcript abundance
of the enzymes of GSH synthesis, but does not affect
GSH concentration (Xiang and Oliver, 1998). Pathogeninduced increases in glutathione in the cells surrounding
the site of attack could have two possible roles in defence.
First, they would increase protection from excessive
damage caused by the accumulation of AOS during the
oxidative burst, which could occur chemically or through
increased substrate availability for enzymes such as
GSTs and GPXs. Second, changes in the redox state
and concentration of glutathione may be an essential
secondary messenger mediating the signalling effects of
hydrogen peroxide (Foyer et al., 1997; May et al., 1998a).
Crucial to signal transduction processes associated with
defence responses appears to be the interaction between
SA, H2O2 and glutathione (Rao and Davis, 1999). There
is also evidence that glutathione is involved in the
regulation of cell division (May et al., 1998a). In plants,
as in animals, cell growth and death responses appear to
be coupled. Morphogenesis involves differential cell division and cell expansion in response to positional cues
generated during development by cell-to-cell communication. At maturity, tissue homeostasis can be influenced by
the action of growth regulators, which act singly or in
combination. Growth and development are also influenced by cell death, which is required for processes such
as the formation of tracheary elements, the release of
mature pollen, the selective elimination of organs during
embryogenesis and, in some species, during flower
development. Damaged cells that are not eliminated via
programmed cell death can proliferate and form tumours.
Glutathione is necessary for the cell to enter the G1 phase,
the pre-mitotic phase of the cell cycle in which the cell
is capable of responding to extracellular stimuli that
determine whether it will enter the S phase, or enter
quiescence, differentiation or death (May et al., 1998a).
Like GSH, GSSG may initiate or potentiate signalling
cascades. GSSG can regulate GR expression as the pea
GR gene contains a putative GSSG binding site (Creissen
et al., 1992). A second, potentially more important
mechanism of GSSG action, involves the spontaneous
oxidation of protein sulphydryl groups to form mixed
1300
Noctor et al.
disulphides, a reaction termed thiolation. While disulphide bonds form spontaneously through a chemical
reaction, re-reduction in vivo requires the intervention of
a protein (e.g. GR, thioredoxin, protein-disulphide isomerase). Hence, some disulphide bonds are transient,
but others can be very long-lived. Reversible disulphide
bond formation has long been recognized as an important
mechanism of modulating protein activity. It has only
recently, however, been shown that this mechanism could
also be a crucial initial signalling event. The formation of
such intramolecular disulphide bonds within proteins
alters their configuration and biological activity (Demple,
1998). Reversible protein thiolation protects essential
thiol groups on key proteins from irreversible inactivation
during oxidative stress and also plays an important
regulatory role in controlling metabolism, protein turnover and gene transcription (Foyer and Noctor, 2001).
There are many examples of proteins that undergo thiolation in animals, but relatively few have been described
in plants. Thiolation has been found, for example, to
activate microsomal GSTs (Dafré et al., 1996), while
thiolation of proteins such as phosphotyrosine-specific
protein phosphatases may also mediate signal transduction pathways that initiate key stress responses
(Fordham-Skelton et al., 1999).
Thiolation may be a particularly important phenomenon in seed development. Some key seed proteins such
as acyl carrier protein are known to be thiolated in the
latter stages of seed development and dethiolated during
imbibition (Butt and Ohlrogge, 1991). Dry seeds often
have a higher glutathione content than other tissues
(Klapheck, 1988; Kranner and Grill, 1996) but much of
the pool is present as GSSG. The seed GSHuGSSG ratio
not only controls protein function but is considered to
regulate protein synthesis as well, in a manner similar
to that observed in mammalian cells (Kranner and Grill,
1996), since low GSHuGSSG ratios block protein synthesis and prevent germination (Fahey et al., 1980).
As soon as germination starts GSSG and protein-thiols
are re-reduced and protein synthesis and function are
re-established. Thiolation of proteins in the dry seeds
could have three functions: firstly, if oxidation of critical
cysteine residues marks proteins for degradation, thiolation will protect both the protein and glutathione from
degradation; secondly, it could modulate protein activity
by interfering with protein cysteine residues; thirdly, it
could be involved in signal transduction associated with
the quiescent state. In addition, low GSHuGSSG ratios
(like low ascorbateuDHA ratios; de Pinto et al., 1999)
block or delay cell division.
Conclusions
The concentration and redox state of intracellular
glutathione pools depends on the complex interplay of
numerous factors. Glutathione redox state is remarkably
constant, but extreme oxidative stress leads to oxidation
of the pool, as observed during ozone exposure or
pathogen attack or in plants with low CAT activities
(Smith et al., 1984; Sen Gupta et al., 1991; Willekens
et al., 1997; Vanacker et al., 2000; Noctor et al., 2001).
Oxidation in these cases is accompanied by increases in
total glutathione. During drought, however, oxidation of
glutathione is not always accompanied by increases in
the pool size (Smirnoff, 1993). The redox state of glutathione will depend on the balance between oxidative
processes and the in vivo GR activity. GR activity may
well reflect reductant supply as much as enzyme capacity,
particularly, perhaps, outside the chloroplast. Interestingly, however, chloroplastic overexpression of either
plant or bacterial GR can increase the leaf GSH:GSSG
ratio and mitigate damage due to certain stresses (Foyer
and Noctor, 2001). Furthermore, several studies have
shown a correlation between GR activity and the absolute
size of the glutathione pool.
The operation of translocators may minimize intercompartmental fluctuations in glutathione concentration
and redox state, and allow different compartments to
co-operate in glutathione synthesis, redox turnover and
degradation. The results presented here on uptake of
GSH into the chloroplast are preliminary, and an
understanding of the influence of chloroplast envelope
transporters will require further characterization of this
process. Transport of other thiols such as Cys and c-EC
is worthy of investigation, particularly in view of the
differential intracellular distribution of c-ECS and
GSH-S observed in tissues such as young wheat leaves.
Another important question is the origin of mitochondrial glutathione and the possibility of a transporter
located on the inner mitochondrial membrane. Identification of genes encoding plant transporters will be
facilitated by the recent cloning of a yeast glutathione
transporter (Bourbouloux et al., 2000).
Acknowledgements
We thank Serge Delrot for discussion, Xiao-Li He for technical
assistance, Julian Coleman for the kind loan of a Beckman
microfuge, and Wacker Chemie GmbH (Munich, Germany) for
the gift of silicone oil. This work was supported by CONICET
Argentina, the European Union, and the UK Biotechnology and
Biosciences Research Council.
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Interactions between biosynthesis, compartmentation and transport

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